1. Field of the Invention
The present invention relates to a method and system that creates a self-contained culture environment, and more particularly to a cell culture system incorporating a disposable cultureware module and a reusable compact instrumentation base device that is capable of expanding cells including primary cells and cell lines as well as patient-specific cells or cells lines in an automated, contaminant-free manner.
2. Description of the Related Art
The anticipated growth of personalized medicine will require new paradigms for the manufacture of therapies tailored to the needs of individual patients. The greatest challenge is expected to come in the area of cell based therapies, especially when such therapies are autologous in nature. In such cases each cell or cell based product will need to be manufactured from scratch for each patient. Manual methods for mammalian cell culture, by their nature, are prone to technician error or inconsistency leading to differences between supposed identical cultures. This becomes especially evident as more and more autologous cells are expanded for personalized therapies. Patient-specific cells, or proteins, are subject to variation, especially when scaled beyond levels that can be managed efficiently with manual methods.
In addition to being labor intensive, the stringent requirements for segregation of each patient's materials from that of every other patient will mean that manufacturing facilities will be large and complex, containing a multitude of isolation suites each with its own equipment (incubators, tissue culture hoods, centrifuges) that can be used for only one patient at a time. Because each patient's therapy is a new and unique product, patient specific manufacturing will also be labor intensive, requiring not just direct manufacturing personnel but also disproportionately increased manpower for quality assurance and quality control functions.
Moreover, conventional approaches and tools for manufacturing cells or cell based products typically involve numerous manual manipulations that are subject to variations even when conducted by skilled technicians. When used at the scale needed to manufacture hundreds or thousands of different cells, cell lines and patient specific cell based therapies, the variability, error or contamination rate may become unacceptable for commercial processes.
Small quantities of secreted product are produced in a number of different ways. I-flasks, roller bottles, stirred bottles or cell bags are manual methods using incubators or warm-rooms to provide environments for cell growth and production. These methods are very labor intensive, subject to mistakes and difficult for large scale production.
Another method, ascites production, uses a host animal (usually a mouse) where the peritoneum is injected with the cells that express the product and are parasitically grown and maintained. The animals are sacrificed and the peritoneal fluid with the product is collected. This method is also very labor intensive, difficult for large scale production and objectionable because of the use of animals. Another method is to inoculate and grow the cells in a small stirred tank or bioreactor or bag-type chamber. The tank provides the environmental and metabolic needs and the cell secretions are allowed to accumulate. This method is costly in terms of facility support in order to do a large number of unique cells and produces product at low concentration.
Another method is to use a bioreactor (hollow fiber, ceramic matrix, fluidizer bed, etc) in lieu of the stirred tank. This can bring facilities costs down and increases product concentration. Biovest International of Coon Rapids, Minn., has or had instruments using these technologies—hollow fiber, ceramic matrix, fluidized bed and stirred tanks.
Cell culturing devices or cultureware for culturing cells in vitro are known. As disclosed in U.S. Pat. No. 4,804,628, the entirety of which is hereby incorporated by reference, a hollow fiber culture device includes a plurality of hollow fiber membranes. Medium containing oxygen, nutrients, and other chemical stimuli is transported through the lumen of the hollow fiber membranes or capillaries and diffuses through the walls thereof into an extracapillary (EC) space between the membranes and the shell of the cartridge containing the hollow fibers. The cells that are to be maintained collect in the extracapillary space. Metabolic wastes are removed from the bioreactor. The cells or cell products can be harvested from the device.
Known EC reservoirs have typically been rigid. They are a pressure vessel and therefore require a sealed compartment with tubing ports adding to costs. A gas, typically air, is introduced through a sterile barrier, generally a membrane filter, to control pressure in the vessel. Fluid level control has been limited to ultrasonic, conductive or optical trip points, or by a load cell measuring the weight of the fluid. Reservoirs are expensive and difficult to manufacture. There is limited EC fluid level measurement accuracy—ultrasonic, conductive or optical monitoring of fluid levels are commonly fouled by cell debris in the reservoir. Alternatively, load cells are not a rugged design for reliable fluid level sensing.
Another problem with the prior art systems is the inability to control lactate and sense pH in the system. One prior art method takes samples of the culture medium and analyzes it using an off-line analyzer. The operator adjusts the perfusion medium rate based on values obtained to maintain the lactate concentration at the level desired. The operator must attempt to predict future lactate levels when adjusting media feed rates. This is labor intensive, presents potential breech of sterility, and the level of lactate control accuracy is dependent on operator skill.
Another method is to connect an automated sampler/analyzer to periodically withdraw sample of the culture media, analyze it and provide feedback for a media feed controller. This method requires additional equipment and increases the risk of sterility breech.
Yet another method is to use an invasive lactate sensor to directly read the lactate level and provide feedback for a media feed controller. In line lactate sensors need to be sterilizable, biocompatible, typically have low reliability and need periodic maintenance.
These methodologies rely on costly, labor intensive off-line sampling and analysis or additional equipment to interface with the instrument or require the addition of a lactate probe and electronics to the culture.
Disposable cultureware generally cannot be autoclaved, so a pH sensor is historically sterilized separately and then added to the cultureware. However, adding the probe risks compromising the sterility of the cultureware. Probe addition is performed in a sterile environment (laminar flow hood) and increases the manpower needed.
The previous methodologies that utilize off-line sampling are subject to contamination problems and depend on the skill of the operator in predicting future lactate levels and influence of media dilution rate. Sampling equipment need interfacing to the culture fluidic circuit, an interface for the feedback signal and periodic calibration of the probes used for sampling. The lactate probe requires interface with the fluid circuit, a method for sterilization or a sterile barrier, interface electronics to convert the probe signal to a useful feedback and a method to calibrate in the fluid circuit.
Preparing the system to start the cell culture is also very labor intensive. The cultureware must be assembled and sterilized or probes must be prepared, sterilized and aseptically inserted into the pre-sterilized portion of the cultureware. The cultureware assembly is then loaded onto the instrument. A series of manual operations are needed to check the integrity of the assembly, introduce fluid into the cultureware flow path, flush the toxic residuals (e.g. surfactants) from the cultureware, start the cultureware in a pre-inoculation mode, introduce factors into the flow path getting it ready for the cells, inoculating the cells into the bioreactor and starting the run (growth of the cell mass and eventual harvest of product).
Two methods are generally used for sterilization. One method places an electrode in a holder, steam sterilizes the assembly (probe) and then aseptically inserts the probe into the pre-sterilized cultureware. The second method involves placing a non-sterile probe into a holder and then using steam to sterilize the electrode in place, referred to as steam in place. Both methods are labor intensive, prone to failure and the procedures need to be validated.
Other methods exist which are less common. Cold sterilants can be used to sterilize the holder and electrode before aseptic insertion. A permeable membrane can be used to isolate the non-sterile probe from the sterile fluid being sensed. A holder with the membrane is placed in the fluid path, either before sterilization or after if the holder and membrane is sterilized separately, and then the sensor is placed against or close to the membrane and the fluid on both sides of the membrane is assumed to be equilibrated.
Glass electrodes have not been included with the cultureware in the past because it was unknown if the probes could survive EtO sterilization and being stored dry. Filled glass electrodes are normally stored hydrated in a liquid buffer.
Each unique cell or cell line must be cultured, cell products harvested and purified separately. In order to do a large number of unique cells or cell lines, a considerable number of instruments would be needed. If application of the cells or products for therapeutic purposes is contemplated strict segregation of each cell production process would be required. Consequently, compactness of the design and the amount of ancillary support resources needed will become an important facilities issue. Moreover the systems currently available are general purpose in nature and require considerable time from trained operators to setup, load, flush, inoculate, run, harvest and unload. Each step usually requires manual documentation.
Moreover, production tracking mandates generation of a batch record for each cell culture run. Historically this is done with a paper-based system and relies on the operator inputting the information. This is labor intensive and subject to errors.
Current purification techniques also involve cleaning and reuse of components. This requires Standard Operational Procedures (SOPS) to be written and the cleaning and reuse process to be validated. This is a time intensive activity.
Accordingly, there is a need for a system and method whereby cells and/or cell products can be cultured in a fully automated, rapid and sterile manner.